In nature, dynamic interactions between enzymes play a crucial role in defining cellular metabolism. By controlling the spatial and temporal organization of these supramolecular complexes called metabolons, natural metabolism can be tuned in a highly dynamic manner. Here, we repurpose the CRISPR–Cas6 family proteins as a synthetic strategy to create dynamic metabolons by combining the ease of RNA processing and the predictability of RNA hybridization for protein assembly. By disturbing RNA–RNA networks using toehold-mediated strand displacement reactions, on-demand assembly and disassembly are achieved using both synthetic RNA triggers and mCherry messenger RNA. Both direct and ‘Turn-On’ assembly of the pathway enzymes tryptophan-2-monooxygenase and indoleacetamide hydrolase can enhance indole-3-acetic acid production by up to ninefold. Even multimeric enzymes can be assembled to improve malate production by threefold. By interfacing with endogenous mRNAs, more complex metabolons may be constructed, resulting in a self-responsive metabolic machinery capable of adapting to changing cellular demand.
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Sequences of all constructs studied are included in the Supplementary Information. Source data are provided with this paper. Additional data are available from the corresponding author upon reasonable request.
Kim, B., Du, J., Eriksen, D. T. & Zhao, H. Combinatorial design of a highly efficient xylose-utilizing pathway in Saccharomyces cerevisiae for the production of cellulosic biofuels. Appl. Environ. Microbiol. 79, 931–941 (2013).
Da Silva, N. A. & Srikrishnan, S. Introduction and expression of genes for metabolic engineering applications in Saccharomyces cerevisiae. FEMS Yeast Res. 12, 197–214 (2012).
Westfall, P. J. et al. Production of amorphadiene in yeast, and its conversion to dihydroartemisinic acid, precursor to the antimalarial agent artemisinin. Proc. Natl Acad. Sci. USA 109, E111–E118 (2012).
Pitera, D. J., Paddon, C. J., Newman, J. D. & Keasling, J. D. Balancing a heterologous mevalonate pathway for improved isoprenoid production in Escherichia coli. Metab. Eng. 9, 193–207 (2007).
Li, N., Zeng, W., Xu, S. & Zhou, J. Toward fine-tuned metabolic networks in industrial microorganisms. Synth. Syst. Biotechnol. 5, 81–91 (2020).
Ferreira, R. et al. Model-assisted fine-tuning of central carbon metabolism in yeast through dCas9-based regulation. ACS Synth. Biol. 8, 2457–2463 (2019).
Toya, Y. & Shimizu, H. Flux controlling technology for central carbon metabolism for efficient microbial bio-production. Curr. Opin. Biotechnol. 64, 169–174 (2020).
Tai, M. & Stephanopoulos, G. Engineering the push and pull of lipid biosynthesis in oleaginous yeast Yarrowia lipolytica for biofuel production. Metab. Eng. 15, 1–9 (2013).
Lan, E. I. & Liao, J. C. ATP drives direct photosynthetic production of 1-butanol in cyanobacteria. Proc. Natl Acad. Sci. USA 109, 6018–6023 (2012).
Xu, P., Li, L., Zhang, F., Stephanopoulos, G. & Koffas, M. Improving fatty acids production by engineering dynamic pathway regulation and metabolic control. Proc. Natl Acad. Sci. USA 111, 11299–11304 (2014).
Zhang, F., Carothers, J. M. & Keasling, J. D. Design of a dynamic sensor-regulator system for production of chemicals and fuels derived from fatty acids. Nat. Biotech. 30, 354–359 (2012).
Win, M. N. & Smolke, C. D. A modular and extensible RNA-based gene-regulatory platform for engineering cellular function. Proc. Natl Acad. Sci. USA 104, 14283–14288 (2007).
Venayak, N., Anesiadis, N., Cluett, W. R. & Mahadevan, R. Engineering metabolism through dynamic control. Curr. Opin. Biotechnol. 34, 142–152 (2015).
Agapakis, C. M., Boyle, P. M. & Silver, P. A. Natural strategies for the spatial optimization of metabolism in synthetic biology. Nat. Chem. Biol. 8, 527–535 (2012).
Menard, L., Maughan, D. & Vigoreaux, J. The structural and functional coordination of glycolytic enzymes in muscle: evidence of a metabolon? Biology 3, 623–644 (2014).
Jørgensen, K. et al. Metabolon formation and metabolic channeling in the biosynthesis of plant natural products. Curr. Opin. Plant Biol. 8, 280–291 (2005).
Møller, B. L. Dynamic metabolons. Science 330, 1328–1329 (2010).
Laursen, T., Møller, B. L. & Bassard, J.-E. Plasticity of specialized metabolism as mediated by dynamic metabolons. Trends Plant Sci. 20, 20–32 (2015).
Dueber, J. E. et al. Synthetic protein scaffolds provide modular control over metabolic flux. Nat. Biotech. 27, 753–759 (2009).
Conrado, R. J., Varner, J. D. & DeLisa, M. P. Engineering the spatial organization of metabolic enzymes: mimicking nature’s synergy. Curr. Opin. Biotechnol. 19, 492–499 (2008).
Chen, R. et al. Biomolecular scaffolds for enhanced signaling and catalytic efficiency. Curr. Opin. Biotechnol. 28, 59–68 (2014).
Berckman, E. A. & Chen, W. Exploiting dCas9 fusion proteins for dynamic assembly of synthetic metabolons. Chem. Commun. 55, 8219–8222 (2019).
Zhao, E. M. et al. Light-based control of metabolic flux through assembly of synthetic organelles. Nat. Chem. Biol. 15, 589–597 (2019).
Chen, R. P., Hunt, V. M., Mitkas, A. A., Siu, K.-H. & Chen, W. Controlling metabolic flux by toehold-mediated strand displacement. Curr. Opin. Biotechnol. 66, 150–157 (2020).
Sachdeva, G., Garg, A., Godding, D., Way, J. C. & Silver, P. A. In vivo co-localization of enzymes on RNA scaffolds increases metabolic production in a geometrically dependent manner. Nucleic Acids Res. 42, 9493–9503 (2014).
Delebecque, C. J., Lindner, A. B., Silver, P. A. & Aldaye, F. A. Organization of intracellular reactions with rationally designed RNA assemblies. Science 333, 470–474 (2011).
Siu, K. H. & Chen, W. Riboregulated toehold-gated gRNA for programmable CRISPR–Cas9 function. Nat. Chem. Biol. 15, 217–220 (2019).
Green, A. A., Silver, P. A., Collins, J. J. & Yin, P. Toehold switches: de-novo-designed regulators of gene expression. Cell 159, 925–939 (2014).
Chen, R. P., Blackstock, D., Sun, Q. & Chen, W. Dynamic protein assembly by programmable DNA strand displacement. Nat. Chem. 10, 474–481 (2018).
Carte, J., Wang, R., Li, H., Terns, R. M. & Terns, M. P. Cas6 is an endoribonuclease that generates guide RNAs for invader defense in prokaryotes. Genes Dev. 22, 3489–3496 (2008).
Niewoehner, O., Jinek, M. & Doudna, J. A. Evolution of CRISPR RNA recognition and processing by Cas6 endonucleases. Nucleic Acids Res. 42, 1341–1353 (2014).
Plagens, A. et al. In vitro assembly and activity of an archaeal CRISPR–Cas type I-A cascade interference complex. Nucleic Acids Res. 42, 5125–5138 (2014).
van der Oost, J., Westra, E. R., Jackson, R. N. & Wiedenheft, B. Unravelling the structural and mechanistic basis of CRISPR–Cas systems. Nat. Rev. Microbiol. 12, 479–492 (2014).
Sokolowski, R. D., Graham, S. & White, M. F. Cas6 specificity and CRISPR RNA loading in a complex CRISPR–Cas system. Nucleic Acids Res. 42, 6532–6541 (2014).
Sternberg, S. H., Haurwitz, R. E. & Doudna, J. A. Mechanism of substrate selection by a highly specific CRISPR endoribonuclease. RNA 18, 661–672 (2012).
Makarova, K. S. et al. Evolution and classification of the CRISPR–Cas systems. Nat. Rev. Microbiol. 9, 467–477 (2011).
Jore, M. M. et al. Structural basis for CRISPR RNA-guided DNA recognition by Cascade. Nat. Struct. Mol. Biol. 18, 529–536 (2011).
Hsu, P. D., Lander, E. S. & Zhang, F. Development and applications of CRISPR–Cas9 for genome engineering. Cell 157, 1262–1278 (2014).
Hall, M. P. et al. Engineered luciferase reporter from a deep sea shrimp utilizing a novel imidazopyrazinone substrate. ACS Chem. Biol. 7, 1848–1857 (2012).
Zadeh, J. N. et al. NUPACK: analysis and design of nucleic acid systems. J. Comput. Chem. 32, 170–173 (2011).
Meng, H. et al. Quantitative design of regulatory elements based on high-precision strength prediction using artificial neural network. PLoS ONE 8, e60288 (2013).
Zhu, L.-y et al. Spatial organization of heterologous metabolic system in vivo based on TALE. Sci. Rep. 6, 26065 (2016).
Ellis, G. A. et al. Artificial multienzyme scaffolds: pursuing in vitro substrate channeling with an overview of current progress. ACS Catal. 9, 10812–10869 (2019).
Škerlová, J., Berndtsson, J., Nolte, H., Ott, M. & Stenmark, P. Structure of the native pyruvate dehydrogenase complex reveals the mechanism of substrate insertion. Nat. Commun. 12, 5277 (2021).
Britton, K. L. et al. The structure and domain organization of Escherichia coli isocitrate lyase. Acta Crystallogr. D 57, 1209–1218 (2001).
Srinivas, N. et al. On the biophysics and kinetics of toehold-mediated DNA strand displacement. Nucleic Acids Res. 41, 10641–10658 (2013).
Charubin, K. & Papoutsakis, E. T. Direct cell-to-cell exchange of matter in a synthetic Clostridium syntrophy enables CO2 fixation, superior metabolite yields, and an expanded metabolic space. Metab. Eng. 52, 9–19 (2019).
This work was supported by grants from NSF (CBET1803008, MCB1817675 and MCB2013991).
A.A.M. and W.C. have filed a patent application (‘Dynamic Control of Colocalization of Proteins’, US patent application 16/751,793) describing the use of orthogonal Cas6 proteins for enzyme assembly and chemical synthesis.
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Extended Data Fig. 1 In vitro Cas6-mediated split Nluc assembly.
a) The assembly components, Csy4-LgBit, Csy4-SmBit and small RNAs A, B, and C, were generated through protein expression and in vitro RNA transcription. b) Production of Cas6 fusions. 7.5% SDS Acrylamide gel indicating full-length expression and purification of the novel fusion proteins Csy4-LgBit and Csy4-SmBit. The full length of the proteins are indicated with around on the bands. Legend is as follows: Sol: soluble fraction, FT: Flow through, W: Wash, E1: Elution fraction 1, E2, Elution fraction 2. c) Different in vitro components were mixed together and luminescence was assayed after 15 minutes of incubation. The two fusion proteins together exhibited minimal background activity as compared to the blank (RLU = 6). A 3-fold increase in luminescence was observed (Lg-A + Sm-B) when the RNA scaffolds added are complementary, while no increase was observed when the two RNA scaffolds do not have complementarity (Lg-A + Sm-C). Results are presented as mean ± s.d. of three biological replicates. d) 10% Urea-PAGE acrylamide gel for confirmation of in vitro transcript products. Products A, B, and C, all should have a full length of around 60 nt. 60 nt is identified on the gel with the red arrow on the gel.
Extended Data Fig. 2 In vivo Cas6-mediated split Nluc assembly.
(a) Original plasmid configuration for in vivo expression of the scaffold system. The p15a plasmid contains a tetracycline inducible promoter, under which the scaffold RNA is placed as well as an arabinose inducible promoter under which the protein Csy4-LgBit-his6x was placed. The pET29b plasmid contains an IPTG inducible promoter under which the Cse3-SmBit-his6x protein was placed. The plasmids were cotransformed in E. coli to assess initial scaffold assembly. This scaffold configuration was used for the experiment in Fig. 1e. (b) Western Blot probing for histidine tags of samples in the experiment in Fig. 1e. The location of each protein is denoted with an arrow on the left of the Western Blot. Each lane represents a different induction condition. (c) Optimized modular expression system. The two fusion proteins were combined onto one plasmid which was placed on the p15a plasmid under the pLtetO-1 promoter. The pET29b plasmid contains the scaffold RNA sequence under control of the pLlacO-1 promoter. Both plasmids were designed to be modular, having unique cut sites around each DNA sequence of interest, which can allow users to easily introduce new RNA scaffold sequences and/or fusion proteins. This plasmid combination was used for the experiment in Fig. 1f. (d) Co-immunoprecipitation schematic illustrating that upon expression of the scaffold RNA, the two fusion proteins are tethered together via RNA interactions and can both be pulled down via an immunoprecipitation using an anti-his antibody. Results of the Western Blot performed on the coimmunoprecipitation elution show successful pulldown of the Cse3-SmBit upon probing with an anti-his mouse antibody. All visible bands on the Western Blot are labelled accordingly.
Extended Data Fig. 3 Effect of longer 5’ RNA handles on split Nluc assembly.
a) Table depicting the characteristics of the three scaffold RNA sequences tested. They are the same overall, with the only difference being the length of the hybridization region. The lengths tested were 13, 18, and 26 nt which correspond respectively to a cis, trans, and cis orientation of the two proteins when they are scaffolded. The number of RNA turns are also highlighted on the table. b) Fold increase in luminescence 8 hours post protein and RNA induction for each of the three different RNA scaffold constructs. Fold increase was calculated as the luminescence ratio of: (proteins + scaffold RNA induction)/ (proteins only induction). Induction conditions were kept the same in all three cases and experiments were performed in parallel. Luminescence results indicate that both proximity of proteins as well as orientation play a role on the assembly of the functional Nluc. Results are presented as mean ± s.d. of three biological replicates.
Extended Data Fig. 4 Improved protein disassembly by mCherry mRNA with media exchange.
Before mCherry mRNA induction via arabinose addition (blue line), the media of the cell culture was exchanged to fresh media without any IPTG or aTc (only arabinose). The luminescence of the sample with trigger expression (blue) drops to 60% of that of the sample with no trigger expression 1 hour post trigger induction and 3 hours post trigger induction it is down to 30%. This result illustrates that the slow reaction observed in Fig. 2d is due to the continued expression of the scaffold RNA due to the presence of IPTG in the medium. Results are presented as mean ± the range of two biological replicates.
Extended Data Fig. 5 Dual plasmid system for dynamic protein assembly.
Expression system for the cycling scaffold architecture. The chloramphenicol resistance plasmid contains the scaffold RNA under pLtetO promoter and the protein operon under the J23113 constitutive promoter system. The two triggers are on the ampicillin resistance plasmid, with trigger 1 under control of the pLlacO-1 promoter and trigger 2 under the pBAD promoter.
Extended Data Fig. 6 Raw luminescence data for the Turn-ON system.
(a) Raw Luminescence data for the direct assembly indicating that there only when the scaffold and proteins are induced (orange) is there a high amount of luminescent signal. (b) Raw luminescence data for the turn ON assembly system. RT: Regular trigger, ScThT: Scrambled toehold trigger, FSc: Fully scrambled trigger. In this case the only conditions upon which any observable increase in luminescence is observed is upon induction of one of the three trigger alongside the proteins (orange, green, yellow).
Extended Data Fig. 7 Expression system for indole-3-acetic acid synthesis.
(a) Plasmid structure for indole-3-acetic acid using direct scaffold assembly. The system is the same as the on in Supplementary Fig. 2(a) except the SmBit and LgBit have now been replaced with the enzymes IaaH and IaaM. The unique restriction sites allowed for easy substitution of the new fusion partners with traditional subcloning techniques. (b) Plasmid structure for indol-3-acetic acid using turn ON scaffold assembly. In this system, the scaffold RNA plasmid has been modified, such that the two separate hairpins of the turn ON system are constitutively expressed via the J23100 synthetic promoter, and the trigger sequence is under control of the pLlacO-1 promoter.
Extended Data Fig. 8 Expression system for enhanced malate synthesis.
(a) Plasmid structure for malate enhancement using direct scaffold assembly. The system is the same as the on in Supplementary Fig. 7(a) except the IaaH and IaaM have now been replaced with the enzymes AceA and AceE. The unique restriction sites allowed for easy substitution of the new fusion partners with traditional subcloning techniques. (b) Plasmid structure for malate enhancement using turn ON scaffold assembly. In this system, the scaffold RNA plasmid has been modified, such that the two separate hairpins of the turn ON system are constitutively expressed via the J23100 synthetic promoter, and the trigger sequence is under control of the pLlacO-1 promoter.
Supplementary Tables 1–6.
Source Data Fig. 1
Raw luminescence data for in vivo split Nluc assembly and statistical analysis for the experiments.
Source Data Fig. 2
Luminescence and fluorescence data for Turn-Off assembly using either a synthetic trigger or mCherry mRNA and statistical analysis for the experiments.
Source Data Fig. 3
Raw luminescence data for recycle assembly and statistical analysis for the experiments.
Source Data Fig. 4
Raw luminescence data and statistical analysis for the Turn-On assembly experiments.
Source Data Fig. 5
IAA production levels and statistical analysis for the experiments.
Source Data Fig. 6
Malate production levels and statistical analysis for the experiments.
Source Data Extended Data Fig. 1
Raw luminescence data for in vitro split Nluc assembly and statistical analysis for the experiments.
Source Data Extended Data Fig. 2
Uncropped pictures for the SDS gel and western blot for the immunoprecipitation experiment.
Source Data Extended Data Fig. 3
Raw luminescence data for effect of hybridization length on split Nluc assembly and statistical analysis for the experiments.
Source Data Extended Data Fig. 4
Raw luminescence data for split Nluc disassembly using mCherry by medium exchange and statistical analysis for the experiments.
Source Data Extended Data Fig. 6
Raw luminescence data for split Nluc assembly using the Turn-On system and statistical analysis for the experiments.
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Mitkas, A.A., Valverde, M. & Chen, W. Dynamic modulation of enzyme activity by synthetic CRISPR–Cas6 endonucleases. Nat Chem Biol 18, 492–500 (2022). https://doi.org/10.1038/s41589-022-01005-7
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